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Originally published In Press as doi:10.1074/jbc.M506995200 on August 29, 2005

J. Biol. Chem., Vol. 280, Issue 45, 37400-37407, November 11, 2005
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Coordinate Control and Selective Expression of the Full Complement of Replication-dependent Histone H4 Genes in Normal and Cancer Cells*

William F. Holmes{ddagger}1, Corey D. Braastad{ddagger}1, Partha Mitra{ddagger}, Cornelia Hampe§, Detlef Doenecke§, Werner Albig§, Janet L. Stein{ddagger}, Andre J. van Wijnen{ddagger}, and Gary S. Stein{ddagger}2

From the {ddagger}Department of Cell Biology and Cancer Center, University of Massachusetts Medical School, Worcester, Massachusetts 01655 and the §Institute for Biochemistry and Molecular Cell Biology, University of Göttingen, Humboldtallee 23, 37073 Göttingen, Germany

Received for publication, June 27, 2005 , and in revised form, August 25, 2005.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The replication of eukaryotic genomes necessitates the coordination of histone biosynthesis with DNA replication at the onset of S phase. The multiple histone H4 genes encode identical proteins, but their regulatory sequences differ. The contributions of these individual genes to histone H4 mRNA expression have not been described. We have determined, by real-time quantitative PCR and RNase protection, that the human histone H4 genes are not equally expressed and that a subset contributes disproportionately to the total pool of H4 mRNA. Differences in histone H4 gene expression can be attributed to observed unequal activities of the H4 gene promoters, which exhibit variations in gene regulatory elements. The overall expression pattern of the histone H4 gene complement is similar in normal and cancer cells. However, H4 genes that are moderately expressed in normal cells are sporadically silenced in tumor cells with compensation of expression by other H4 gene copies. Chromatin immunoprecipitation analyses and in vitro DNA binding assays indicated that 11 of the 15 histone H4 genes interact with the cell cycle regulatory histone nuclear factor P, which forms a complex with the cyclin E/CDK2-responsive co-regulator p220NPAT. These 11 H4 genes account for 95% of the histone H4 mRNA pool. We conclude that the cyclin E/CDK2/p220NPAT/histone nuclear factor P signaling pathway is the principal regulator of histone H4 biosynthesis.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Histones have crucial roles in replication, transcription, repair, and recombination (1–3). There is a fundamental requirement for coordinated de novo synthesis of the core histone proteins H2A, H2B, H3, and H4 as well as the linker H1 protein during S phase to package nascent genomic DNA (1, 2). Replication of a complete mammalian genome requires 108 of each of the individual histone proteins. Efficient production of this vast quantity of proteins necessitates that transcription of multiple histone genes at multiple loci be coordinately regulated with the onset and progression of genome replication during the cell cycle (4).

Histone biosynthesis is a unique process involving transcription initiation from compact promoters to form primary transcripts that lack introns and that contain a highly conserved stem-loop structure that forms the 3'-end of the mature non-polyadenylated mRNA (4, 5). Histone genes are organized into clusters, and this organization has persisted throughout the course of evolution from yeast to human (2, 4). The majority of the 74 known and characterized human histone genes is located in two major clusters at chromosomes 6p21 and 1q21, respectively (TABLE ONE) (1, 6–8). It is now known that the human genome contains 15 histone H4 genes that encode identical proteins. H4 genes in lower eukaryotes (e.g. sea urchin and Drosophila) are organized with the other histone gene types (i.e. H2A, H2B, H3, and H1) into units that are tandemly repeated, and all H4 genes in these organisms have virtually identical promoters and coding regions. Although the coding regions of the human histone H4 genes are translated into identical proteins, there is surprising variation in the organization of the proximal promoters. Based on the availability of the complete human genome sequence, it is now possible to definitively assess the expression and regulation of the full complement of histone H4 genes.


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  		TABLE ONE
Human histone H4 genes

H4/f is a pseudogene (7).

 
Previous studies on a limited number of H4 genes have suggested that the expression of many (if not all) of these H4 copies is coordinately controlled during the cell cycle (2, 9–11). The human histone H4/n gene, which is temporally and functionally linked to DNA synthesis, provides a paradigm for cell cycle-dependent coordinate control of gene expression at the G1/S phase transition. This H4 gene is regulated by multiple elements and cognate DNA-binding activities, and one proximal promoter element (Site II) mediates cell cycle-dependent transcription (12–17). A phylogenetically conserved H4 subtype-specific element is typically present in H4 genes within Site II. Site II is not responsive to the E2F class of transcription factors. Thus, Site II cell cycle regulatory mechanisms at the onset of S phase function independently of E2F (2, 15, 18). Three histone nuclear factors (HiNF-M, -D, and -P)3 interact with Site II to mediate cell cycle control of transcription at the onset of S phase (4, 12–24). Recently, HiNF-P has been identified as the protein that, in conjunction with the cyclin E/CDK2-responsive protein p220NPAT, controls H4 gene transcription via binding to its cognate H4 subtype-specific element (25). It is necessary to assess whether all human histone H4 genes are coordinately controlled and whether HiNF-P is the key factor that synchronizes transcription of the histone H4 gene family with DNA replication.

Because of the extensive similarity of the histone H4 sequences, it has been difficult to determine the relative contributions of the 15 individual H4 genes to the total histone H4 mRNA pool by molecular approaches that require hybridization of relatively large nucleic acid probes (i.e. Northern blotting and RNase protection). However, we have circumvented this limitation by applying PCR-based approaches that discriminate between individual histone genes and transcripts. In this study, we used quantitative PCR (qPCR) and chromatin immunoprecipitation (ChIP) to examine the expression and regulation of the 15 human histone H4 genes in normal and tumor-derived cells. One key finding of our study is that the expression levels of the individual genes differ considerably, consistent with variations in H4 promoter organization and activity. Furthermore, our results firmly establish that 11 genes, which account for >95% of histone H4 mRNAs, are coordinately controlled during the cell cycle and are responsive to the cyclin E/CDK2/p220NPAT/HiNF-P signaling pathway.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Lines and Culture Conditions—COS-7, HeLa, HCT116, IMR90, SaOS, and T98G cells were obtained from American Type Culture Collection (Manassas, VA). The COS-7, HeLa, and T98G stock cultures were maintained in Dulbecco's modified Eagle's medium (Invitrogen) supplemented with 10% fetal bovine serum, 2 mM L-glutamine, 100 units/ml penicillin, and 100 µg/ml streptomycin. HCT116 and SaOS cells were maintained in McCoy's 5A medium (Invitrogen) supplemented with 10 or 15% fetal bovine serum, respectively, 2 mM L-glutamine, 100 units/ml penicillin, and 100 µg/ml streptomycin. IMR90 cells were maintained in Eagle's basal medium (Invitrogen) supplemented with 10% fetal bovine serum, 2 mM L-glutamine, 100 units/ml penicillin, and 100 µg/ml streptomycin. All cell lines were split at a ratio of 1:10 when 70% confluent in 100-mm plates (Corning Inc., Corning, NY). Culture conditions were 37 °C in a 98% humidified and 5% CO2 incubator.

Quantitative Reverse Transcription-PCR (qRT-PCR) Analysis of H4 Gene Expression—RNA was extracted from specified cell lines using TRIzol® reagent (Invitrogen) according to the manufacturer's protocol. Purified total RNA was subjected to DNase I digestion, followed by column purification using the DNA Free RNA KitTM (Zymo Research Corp., Orange, CA). Eluted total DNA-free RNA was quantitated by spectrophotometry, and 1 µg was added to a reverse transcription reaction using the iScriptTM cDNA synthesis kit (Bio-Rad) with a mixture of random hexamers and oligo(dT) primers. Varying dilutions of cDNA were used as templates in qPCRs with oligonucleotides specific to the different histone H4 gene 5'-untranslated regions (see TABLE TWO). Relative quantitation was determined using an ABI PRISM 7000 sequence detection system (Applied Biosystems) measuring real-time SYBR Green (Bio-Rad) fluorescence and calculated by the {Delta}{Delta}CT method as described recently (26). Overall efficiencies of qPCR were calculated from the slopes of the standard curves of serial dilutions in steps of 2 (log(2) scale) and found to be nearly identical for each primer set. Expression profiles for H4/a mRNA were extrapolated by comparing qPCRs with H4/a fluorescent minor groove binder probe to qPCRs with both SYBR Green and minor groove binder probes specific for H4/n mRNA.


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  		TABLE TWO
Specific primers for the individual histone H4 transcripts

FAM, 6-carboxyfluorescein; MGB, minor groove binder.

 
RNase Protection Assays (RPAs)—RPAs were performed as described (27). Briefly, 32P-labeled antisense RNA was transcribed in vitro from linearized template DNA using the MAXIscript T7/T3 kit (Ambion, Inc., Austin, TX) in the presence of 3.3 mM [{alpha}-32P]UTP (3000 Ci/mmol, 50 mCi) and 50 mM unlabeled UTP for use in RPAs (RPA II kit from Ambion, Inc.). [{alpha}-32P]UTP-labeled {beta}-actin antisense RNA was added to each hybridization reaction to normalize RNA quantities. Protected RNA fragments were resolved on a denaturing 8 M urea and 5% (v/v) polyacrylamide gel. A 32P-end-labeled Sau3AI digest of pUC19 DNA was used as a size marker. A PhosphorImager (Amersham Biosciences) was used to quantify protected RNA fragments. The relative expression of the individual histone H4 genes was calculated as the ratio of signal intensity standardized by {beta}-actin. To compare RPA data with qRT-PCR data, the relative expression of H4/d was set to a value of 100% (maximum), and the relative expression levels of the other H4 mRNAs were determined as a percent of this maximal level.

Reporter Gene Analysis—In all experiments, promoter activity was measured in whole cell extracts using a luciferase assay kit (Promega Corp., Madison WI), and the results were normalized by cotransfection with 1 µg of pCMV-{beta}-gal. To determine the expression of histone H4 as a function of the HiNF-P·p220NPAT complex, cells were transiently transfected with either 150 ng of wild-type HiNF-P or 300 ng of an expression vector containing wild-type p220NPAT or both with 200 ng of one of the wild-type histone H4 promoter-luciferase reporter constructs. All cDNAs and reporter constructs were mixed with FuGENE 6 (Roche Applied Science) in 100 µl of serum-free medium for 20 min at room temperature and then applied to cells. Cells were incubated overnight with the DNA mixture in a final volume of 2 ml of medium and then harvested 24 h later. Luciferase activity was measured using a Monolight 2010 luminometer (Analytical Luminescence Laboratory, San Diego, CA).

ChIP—Synchronously or asynchronously growing adherent cells were washed twice with ice-cold 1x phosphate-buffered saline (PBS) on the plate. Cells were immediately treated with 5 ml of 1% formaldehyde cross-linker in 1x PBS at room temperature for 10 min with gentle rotation. The cross-linking reaction was quenched by the addition of 5 ml of 0.25 M glycine in 1x PBS at room temperature for 5 min. Cells were then washed twice more with ice-cold 1x PBS, scraped into 5 ml of 1x PBS on ice, and harvested by centrifugation at 1000 x g for 5 min at 4 °C. Cross-linked cell pellets were resuspended in 50 mM Tris-Cl (pH 8.1), 150 mM NaCl, 1% (v/v) Nonidet P-40, and 2x Complete protease inhibitor mixture (Roche Applied Science) and incubated on ice for 20 min. Lysates were then sonicated to an average DNA size of 500–100 bp by agarose gel electrophoresis, and the extracts were cleared by centrifugation at 14,000 x g for 15 min at 4 °C. Cleared extracts were divided into sample and input aliquots to allow subsequent quantitation. Sample aliquots were subjected to primary immunoprecipitation with 2 µg of purified immunoglobulin or 3 µl of crude serum for each appropriate antibody. Following primary antibody incubations, a 0.1 volume of protein A/G-agarose bead slurry (Santa Cruz Biotechnology, Inc., Santa Cruz, CA) was added and allowed to bind for 1–4 h. Immunocomplexes bound to the beads were harvested by centrifugation at 1000 x g for 3 min. The beads were then washed consecutively with the following buffers: a low salt buffer containing 20 mM Tris-Cl (pH 8.1), 150 mM NaCl, 1% Triton X-100, 2 mM EDTA, and 1x Complete protease inhibitor mixture; a high salt buffer containing 20 mM Tris-Cl (pH 8.1), 500 mM NaCl, 1% Triton X-100, and 2 mM EDTA; 10 mM Tris-Cl (pH 8.1), 250 mM LiCl, 1% deoxycholate, 1% Nonidet P-40, and 1 mM EDTA; and three washes with 10 mM Tris-Cl (pH 8.1) and 1 mM EDTA. Crosslinked protein·DNA complexes were eluted from antibodies and beads twice with 100 µl of 1% SDS and 100 mM NaHCO3. The pooled eluates were supplemented with 0.1 volume of 3 M sodium acetate (pH 5.2), and the cross-links were reversed at 65 °C overnight. DNA was then purified via phenol/chloroform extraction and isopropyl alcohol precipitation with 5–20 µg of glycogen carrier. Precipitated DNA was allowed to rehydrate in 10 mM Tris-Cl (pH 8.1), and material ratios between samples and inputs were carefully documented to allow subsequent quantitation of locus immunoprecipitation. ChIP samples were then subjected to qPCR analysis using the ABI PRISM 7000 sequence detection system with locus-specific primers and probes.

Electrophoretic Mobility Shift Assay (EMSA)—In vitro DNA binding of HiNF-P to selected histone H4 genes was analyzed as described (13, 25). The radiolabeled probe we used represents an optimized HiNF-P-binding site that is based on the HiNF-P recognition site in Site II of the H4/n gene (5'-CTT CAG GTT TTC AAT CTG GTC CGA TAC T). The probe was incubated with HiNF-P-enriched nuclear extract from HeLa cells. Competition experiments were carried out with a 50-, 100-, 200-, or 400-fold excess of unlabeled oligonucleotides spanning the analogous regions in the promoters of distinct H4 genes.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
mRNAs of Multiple Human Replication-dependent Histone H4 Genes Are Coordinately Induced in S Phase Cells—We predicted temporal and/or quantitative differences in expression of the 15 human histone H4 genes because of the heterogeneity in 5'-flanking sequences of the individual genes. Therefore, we initially examined the relative levels of expression of distinct H4 mRNAs during the cell cycle. Human T98G glioblastoma cells were serum-deprived for 72 h to induce quiescence and then stimulated to re-enter the cell cycle with serum. Synchronously cycling cells were harvested at 4, 21, and 25 h post-stimulation, corresponding to the G1, S, and G2 phases of the cell cycle, respectively, as determined by fluorescence-activated cell sorter analysis (Fig. 1A). Total RNA was examined by qRT-PCR. The relative contributions of individual histone H4 genes to the total H4 mRNA pool were determined with primer pairs specific to the unique 5'-untranslated region of each H4 gene (TABLE TWO). Primer pairs were tested against dilutions of genomic DNA to verify that the amplification efficiencies were similar. The mRNA levels of each histone H4 gene were up-regulated in S phase between 5- and 20-fold above the levels in G1 and G2 phases (Fig. 1B). This result is the first demonstration that the complement of H4 genes in the human genome is temporally and coordinately controlled at the G1/S phase transition and is consistent with previous studies that examined a limited number of representative human histone H4 genes (9–11, 13, 20, 22, 25, 28).



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  		FIGURE 1.
Cell cycle-dependent coordinate control of histone H4 gene expression. A, fluorescence-activated cell sorter profiles are shown for T98G cells synchronized by serum deprivation and harvested at 4, 21, or 25 h post-serum stimulation, corresponding to the G1, S, and G2 phases, respectively. B, all H4 gene mRNA levels are induced from G1 to early S phase between 5- and 20-fold. In G2 phase cells, all H4 gene mRNA levels are similar to those in phase G1 cells. H4/d and H4/e ({diamondsuit}), as well as H4/n and H4/o ({diamondsuit}), are respective genomic duplicates and cannot be distinguished from one another by qPCR. Therefore, the relative contribution of each of these duplicated genes was estimated as half of the total when the following symbols are used: {diamondsuit} and {diamondsuit}.

 
Our results also indicate that there are differences in the extent to which distinct H4 mRNA species are maximally expressed. We found that the H4/b, H4/d, H4/e, H4/j, H4/n, and H4/o genes account for >76% of the total histone H4 mRNA during S phase. H4/g, H4/h, and H4/i are moderately expressed, together representing 20% of the total histone H4 mRNA, whereas H4/a, H4/c, H4/k, H4/m, and H4/p contribute minimally to the total H4 mRNA pool (<5%). Thus, nine of the histone H4 genes generate the majority of mRNAs that are required to accommodate DNA replication and nascent chromatin assembly.

Differential Regulation of Histone H4 Gene Expression in Normal and Transformed Human Cells—The promoter elements of H4 genes interact with a multiplicity of transcription factors that respond to cell growth-related signaling pathways that may be differentially activated in normal versus tumor cells. Thus, we assessed whether the deregulation of signaling pathways in transformed and tumor cells affects the composition of H4 mRNA pools. We compared the expression patterns of individual human histone H4 genes in fetal liver, fetal colon, and IMR90 (normal diploid lung fibroblast) cells with those in four different tumor-derived human cell lines (i.e. HCT116 colorectal carcinoma cells, T98G glioblastoma cells, SaOS osteosarcoma cells, and HeLa cervical carcinoma cells).

We initially compared the RPA data with the qRT-PCR data in two human carcinoma lines (data not shown). The RPAs and the qRT-PCRs were performed in completely independent experiments. The two assays showed consistent quantitative differences in the relative expression of individual histone H4 genes in asynchronous HeLa and SaOS cell lines. In additional studies, we used qRT-PCR as the primary method to determine the relative contributions of the individual histone H4 genes in various cell lines and tissues.

The most striking result is that the expression levels of the individual H4 genes are substantially different regardless of the cell type and that the relative levels are distinct from the theoretically expected contribution of ~7% per gene copy (expression of 15 genes is 100%) (Fig. 2, A and B, dashed lines). In three normal and four tumor-derived cell types, we found that the five most highly expressed genes (H4/d, H4/e, H4/j, H4/n, and H4/o) contribute the majority of H4 mRNAs to the total pool. The only quantitative difference is that these highly expressed genes contribute a disproportionate percentage of the total H4 mRNAs in tumor-derived cells compared with normal cells. For example, the above five highly expressed genes and six modestly expressed H4 genes (e.g. H4/a, H4/b, H4/c, H4/k, H4/m, and H4/p) contribute 55 and 18% in normal cells, whereas these contributions are 80 and 9% in tumor-derived cells, respectively. These differences can be attributed primarily to undetectable mRNA levels of a subset of H4 genes in tumor-derived cells (i.e. H4/b, H4/c, H4/k, and/or H4/m). These results suggest that modestly expressed H4 genes are silenced in tumor cells with concomitant compensatory responses by other more highly expressed H4 genes.



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  		FIGURE 2.
Expression patterns of the human replication-dependent histone H4 genes in various cell types. Relative histone H4 gene expression profiles were determined by qRT-PCR in two normal fetal tissues and normal diploid-derived cells (A) and in four tumor-derived cells (B). The profiles are presented as percent contribution to total H4 mRNA pools. The dashed lines indicate the theoretical level of contribution if each H4 gene were equally expressed (15 genes = 100%). The relative contribution of each duplicated gene was estimated as half of the total ({diamondsuit} and {diamondsuit}). Error bars represent the S.D. of triplicate experiments. ND, not determined.

 
Basal H4 Promoter Activity Dictates H4 mRNA Accumulation—It is well documented that all histone H4 mRNAs contain a 3'-stem-loop structure (hairpin) that is necessary for 3'-end processing and mRNA stability (8). Because the hairpin sequences are completely conserved among all H4 genes, it is unlikely that the differential expression of H4 gene mRNAs is due to these post-transcriptional mechanisms. Rather, quantitative differences and temporal similarities in the expression of H4 gene copies may reflect the divergence and conservation of distinct 5'-regulatory elements within H4 promoters. Therefore, we tested whether the observed changes in expression of H4 gene copies are related to differences in H4 gene promoter activity. We monitored luciferase reporter gene expression using promoters of all H4 genes located within the two major histone gene superclusters (human chromosomes 1 and 6) in multiple cell types. The promoters with the highest transcriptional activity in normal and tumor-derived cells include H4/d, H4/j, and H4/n and, by inference, also H4/e and H4/o, as their corresponding promoters are essentially duplicates of H4/d and H4/n, respectively (Fig. 3). These are the same H4 genes that exhibit high endogenous levels of expression (Fig. 2). Furthermore, the activities of individual H4 promoters are more similar to the average in normal cells than in tumor-derived cells, mirroring the endogenous H4 mRNA expression patterns (Figs. 2 and 3). Thus, differential expression of H4 genes is transcriptionally mediated, as expected from the divergence in the overall organization of regulatory elements within H4 promoters.

The H4/n gene has been shown to be regulated by a highly conserved cell cycle regulatory domain (Site II) and an auxiliary module (Site I) that augments the transcription rate. Regions analogous to Site II of the H4/n gene are present in the other H4 genes and can easily be aligned (Fig. 4). However, there is considerable heterogeneity in promoter organization beyond Site II. Alignment of the 15 human Site II sequences revealed that there are four H4 genes (i.e. H4/a, H4/c, H4/k, and H4/l) that exhibit clear mismatches with the H4 subtype-specific consensus element, whereas a fifth H4 gene (i.e. H4/m) exhibits a single nucleotide deviation in the TATA box. These five genes generally have lower than average expression and promoter activity in different cell types. This finding is consistent with our previous result that Site II is a positive element that mediates the cell cycle-dependent activation of H4 gene transcription (15, 25).



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  		FIGURE 3.
Basal promoter activity of the individual H4 genes. Shown is the transient transfection of the individual H4 proximal promoters driving luciferase (Luc) expression in normal diploid IMR90 cells as well as in T98G, SaOS, and HCT116 tumor-derived cell lines. Luciferase activities were measured as relative light units (RLU) and normalized with cotransfected pCMV-{beta}-galactosidase ({beta}-gal). The dashed lines indicate the average level of normalized expression of all analyzed H4 reporter constructs. H4/d and H4/e ({diamondsuit}), as well as H4/n and H4/o ({diamondsuit}), are respective promoters and cannot be distinguished, and activity from a single proximal promoter construct is representative for both genes ({diamondsuit} and {diamondsuit}). Error bars represent the S.D. of triplicate experiments. ND, not determined.

 
The principal factor that interacts with Site II of the H4/n gene is HiNF-P (25). Because deviations in the H4 subtype-specific consensus sequence correlate with reduced expression, we examined whether nucleotide changes decrease the level of HiNF-P binding. Competition EMSAs were performed with the HiNF-P consensus oligonucleotide as a probe and increasing amounts of unlabeled Site II competitors spanning a select set of H4 genes. Some of these oligonucleotides contain critical mutations that are predicted to decrease or abolish HiNF-P binding (Fig. 5). Lack of binding is reflected by the absence of competition at a 400-fold molar excess, as has been observed for an H4/n-derived oligonucleotide with critical mutations in the HiNF-P contact points (13). Our results show that the Site II sequences of the H4/a and H4/k genes do not bind HiNF-P, whereas the H4/c element binds very weakly (Fig. 5). In contrast, the Site II sequences of the H4/g and H4/p genes interact, as expected, very efficiently with HiNF-P. Together with previous studies that compared the interactions of HiNF-P with the H4/n and H4/o genes and the H4/d and H4/e genes (13, 25), our studies establish a strong relationship between matches in the Site II/HiNF-P consensus sequence and binding of HiNF-P.

Histone H4 Gene Expression as a Function of Promoter Occupancy—Having established the effects of Site II mutations on HiNF-P binding in vitro, we addressed whether differential regulation of H4 genesis due to differences in RNA polymerase II association and the ability of HiNF-P to bind to H4 loci in vivo. Gene-specific promoter occupancy by these two proteins was assayed in asynchronous T98G cells by ChIP and qPCR. Promoter occupancy was determined by the specific presence of various H4 promoter sequences in immunoprecipitates obtained using antibodies to HiNF-P and RNA polymerase II (Fig. 6).

We found a positive correlation between HiNF-P and RNA polymerase II recruitment to the H4 promoters and the corresponding mRNA levels, although the presence of HiNF-P and/or RNA polymerase II does not guarantee high transcript levels. The genomic promoters of all H4 genes examined, with the exception of H4/a, H4/c, and H4/m, interact with HiNF-P in vivo. The genes that do not interact with HiNF-P are minimally expressed and do not contribute appreciably (<3%) to the overall H4 mRNA pool. This finding supports our previous (25) and present results that HiNF-P occupancy of H4 gene loci is necessary for optimal expression and that HiNF-P is a primary regulator of H4 gene transcription.

Highly Expressed H4 Genes Are Responsive to the Cyclin E/CDK2/p220NPAT/HiNF-P Signaling Pathway—HiNF-P activation of the H4/n gene depends critically on coactivation by the cyclin E/CDK2-responsive p220NPAT protein, and endogenous HiNF-P and p220NPAT levels are limiting for H4/n gene transcription (25). We assessed which of the multiple histone H4 promoters are regulated by this signaling pathway using reporter gene assays in which HiNF-P and p220NPAT were coexpressed (Fig. 7). For example, the H4/n gene is up-regulated 3-fold by HiNF-P or p220NPAT alone (data not shown) and is synergistically activated 10-fold or more when both proteins are coexpressed (Fig. 7), consistent with our previous observations (25). As expected, the H4/a, H4/c, and H4/m genes, which do not bind HiNF-P as determined by EMSA and ChIP-qPCR analysis (Figs. 5 and 6), do not respond to the HiNF-P/p220NPAT signaling pathway in HCT116, T98G, and SaOS cells. H4/c and H4/m genes respond modestly in IMR90 cells, which appear to be the exception. However, the induced activities of these two promoters remain quite low in IMR90 cells, perhaps reflecting an indirect effect of HiNF-P/p220NPAT. More importantly, of the 11 HiNF-P-dependent genes that we analyzed, seven are robustly up-regulated by p220NPAT signaling in all cell types and three in at least two cell types. The 11th gene (H4/g) responds qualitatively, but exhibits very low induced promoter activity. We conclude that all 11 HiNF-P-responsive H4 genes are also co-responsive to p220NPAT. Because these genes contribute to >95% of the total H4 mRNA pool (Fig. 2), it appears that the HiNF-P/p220NPAT signaling pathway is essential for coordinate control of histone H4 gene expression.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
In this study, we have examined the similarities and differences among the 15 human histone H4 genes in cell cycle control, expression in normal and tumor-derived cells, proximal promoter activity, and promoter binding of transcription factors. Our results firmly establish coordinate cell cycle control of the majority of histone H4 gene expression as a result of the cyclin E/CDK2/p220NPAT/HiNF-P signaling pathway. Furthermore, a subset of these genes are sporadically silenced in tumor cell lines in comparison with normal cells. The unequal expression of H4 genes may reflect differences in position within nuclear architecture and/or distinctive promoter element organization. We have observed that histone H4 proximal promoter activities are consistent with the expression levels of endogenous H4 mRNAs. Finally, we have established that all highly expressed H4 genes are regulated by the cell cycle-dependent transcription factor HiNF-P.

Coordinate Induction of H4 Gene Expression during the G1/S Phase Transition—Many previous studies have focused on the cell cycle-dependent expression of total histone mRNAs as determined by northern blot analysis. A subset of these studies included assays capable of distinguishing one or more histone gene copies. However, as these studies were performed before the human genome project was completed, none of the previous findings permitted a complete analysis of the individual contribution of all members of the histone gene family.

It has been well established that the biosynthesis of histone H4 is mediated by multiple functionally expressed H4 genes (2, 9–11). By comprehensive analysis of the expression of the human histone H4 gene family in synchronized cells, one major finding of this study is that all mRNAs derived from the multiple human histone H4 genes are indeed simultaneously up-regulated when cells progress into S phase. Because post-transcriptional mechanisms operating on distinct H4 mRNAs are expected to be identical, this coordinate regulation is directly attributable to transcriptional mechanisms. Although our data now conclusively establish that coordinate regulation of the full complement of histone H4 genes does indeed occur, the results indicate that a surprisingly small number of genes account for the majority of H4 gene expression. We found that the promoters of the histone H4 genes are not regulated in an equivalent manner by cell cycle-driven signaling events. Interestingly, two of the histone H4 genes (H4/n and H4/o) that are most responsive to signals at the G1/S phase transition are recent duplications in the human genome (6). This recent duplication may reflect a preferred preservation and expansion of H4 genes with the requisite regulatory organization to support DNA replication and cell survival.



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  		FIGURE 4.
Alignment of the Site II cell cycle regulatory element in the full complement of human histone H4 genes. Mismatches in Site II, which spans both the H4 subtypespecific consensus element (cons) and the TATA box, are examined in relation to H4 mRNA expression and H4 promoter strength. HiNF-P binds to DNA by interacting with the conserved residues of the H4 subtype-specific cell cycle element to induce transcriptional activation of H4 genes in concert with G1/S phase transition signals (13, 14, 16, 17, 22, 25, 28). The sites of HiNF-P contact within the consensus element are indicated (*). Mismatches with the consensus sequence are indicated by boldface lowercase letters. Redundant nucleotides (nt) are indicated as follows: D = A, G, or T; H = A, C, or T; M = AorC; W = AorT;and Y = C or T. The columns to the right correlate sequence data with H4 gene expression and transcription. H4 mRNA expression data are summarized as follows: +, below average levels in most or all cell lines; ++, intermediate levels in several cell lines; +++, above average expression in all cell lines; ND, not determined. H4 promoter strength data are summarized as follows: –, no significant activity; +, low activity in most or all cell lines; ++, intermediate activity in several cell lines; +++, strong activity in most or all cell lines; ND, not determined.

 



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  		FIGURE 5.
In vitro interactions between HiNF-P and Site II equivalents in selected H4 genes. A, EMSAs were performed with HeLa nuclear protein and a standard radiolabeled HiNF-P-binding site oligonucleotide in the absence (control) or presence of increasing amounts of unlabeled competitor oligonucleotides (50-, 100-, 200-, and 400-fold molar excesses, representing total binding site concentrations of 25, 50, 100, and 200 nM, respectively) spanning variant Site II sequences derived from the genes indicated. The arrowheads indicate two protein·DNA complexes: HiNF-P (P) and an unrelated complex (u). B, shown is a summary of matches of Site II equivalents in the indicated H4 genes (left column: +, 100% match; and –, mismatch) with binding of HiNF-P to Site II in vitro by EMSA (second column; data presented in A) and in vivo by ChIP (third column; data presented in Fig. 6). The fourth column indicates the sensitivity of the distinct H4 genes to treatment with HiNF-P antisense oligonucleotides (+, HiNF-P-dependent and H4 mRNA-decreased; –, not HiNF-P-dependent) as previously documented (28). *, the H4/n and H4/o genes are recent duplicates, and the results are based on the same oligonucleotide; **, the same is true for the H4/d and H4/e genes, and data comparing the H4/n (FO108) and H4/d (HuH4a) genes were published previously under different gene names (13, 25); #, these are exceptions to the rule that HiNF-P binding in vitro predicts in vivo interactions and HiNF-P dependence. (Note there may be uncharacterized HiNF-P sites elsewhere in the H4/k gene, whereas the H4/m gene has a deviant TATA box as shown in Fig. 4.) ND, not determined.

 



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  		FIGURE 6.
Histone H4 gene expression as a function of promoter occupancy by a crucial H4 transcription factor and RNA polymerase II. In T98G cells, ChIP was performed with antibodies to HiNF-P and RNA polymerase II (RNAP II). qPCR for the promoter/5'-region of each histone H4 gene (x axis) was performed on the ChIP samples and compared with the input to establish the percentage of each locus that was immunoprecipitated (y axis = percent input). The relative contribution of each duplicated gene was estimated as half of the total ({diamondsuit} and {diamondsuit}). The inset shows a sample ChIP experiment using the H4/n gene with pertinent controls for specificity. Negative controls included nonspecific IgG and immunoprecipitates from immunoprecipitation with anti-HiNF-P and anti-RNA polymerase II antibodies that was performed in the presence of the corresponding immunogenic peptides. Error bars represent the S.D. of triplicate experiments. Additional negative control experiments included analysis of precipitates with qPCR primers against functionally expressed non-histone H4 genes (positive for RNA polymerase II and negative for HiNF-P) as well as non-expressed genomic segments (negative for both RNA polymerase II and HiNF-P) (data not shown). In all cases, we observed background values for negative controls. The qPCR results shown here were consistent with and more quantitative than the results from gel-based ChIP assays for the H4/n (FO108) gene that we presented previously (25).

 



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  		FIGURE 7.
The majority of H4 gene promoters is synergistically induced by HiNF-P and p220NPAT. Normal and tumor-derived cell lines were transiently cotransfected with the individual H4 proximal promoters with (black bars) and without (white bars) expression vectors for HiNF-P and p220NPAT. -Fold change in expression is indicated for the promoters of H4 genes that have basal activity above average levels (dashed lines in Fig. 2). H4/d and H4/e ({diamondsuit}), as well as H4/n and H4/o ({diamondsuit}), are respective promoters and cannot be distinguished from one another, and activity from a single proximal promoter construct is representative for both genes ({diamondsuit} and {diamondsuit}). Error bars represent the S.D. of triplicate experiments. RLU, relative light units; {beta}-gal, {beta}-galactosidase; ND, not determined.

 
Histone H4 Genes Contribute Differentially to the Replicationdependent Pool of Total Histone H4 mRNA—Due to the dysfunctional regulation of many cell cycle-related factors, carcinoma cells will differentially activate replication-dependent promoter elements of various genes. We have determined that this differential activation extends to histone H4 genes and may not be a function of tissue specificity. Comparing the expression of histone H4 genes in normal cells with that in human carcinoma cells revealed that a subset of individual H4 genes are differentially expressed distinct from any theoretically expected contribution to the total H4 mRNA pool. The disparity in the relative levels of this subset of H4 mRNAs is even greater in carcinoma cell lines, substantiating that aberrant cells have dysfunctional replication-dependent signals.

The Promoters of Histone H4 Genes Exhibit Variations in Promoter Element Organization—The burden of coordinate control of mRNA levels is shared by two main mechanisms, transcriptional induction and increased mRNA stability by 3'-endonucleolytic processing. However, it is well documented that all histone H4 3'-stem-loop structures are completely conserved. Therefore, the variation in the endogenous expression of histone H4 mRNAs in normal and carcinoma cells is directly attributable to the relative activities of the individual H4 gene promoters. Alignment of the H4 subtype-specific cell cycle elements revealed that some residues are necessary for HiNF-P binding and consequently cell cycle coordinate control of H4 expression at the G1/S phase transition. Furthermore, variation in the H4 subtype-specific element sequences causes some H4 genes to be poorly responsive to HiNF-P and results in lower contribution to the total pool of H4 mRNA. Thus, the variation in the contributions of the individual H4 genes is not likely due to mRNA stability, but rather to divergence of promoter elements and/or overall chromatin organization of histone gene clusters.

Highly Expressed H4 Genes Are Responsive to the HiNF-P Signaling Pathway—HiNF-P and RNA polymerase II occupancy of histone H4 promoters does not completely correlate with H4 gene mRNA contribution to the total H4 mRNA pool. Recruitment of RNA polymerase II to low expressing genes may indicate a stalled RNA polymerase II complex. Lack of promoter occupancy by HiNF-P is typical for low expressing H4 genes, albeit that Site II occupancy by HiNF-P does not assure high transcription rates. This finding suggests that HiNF-P is required for high level expression, but insufficient for maximal activation of cell cycle-dependent H4 gene transcription. HiNF-P activation of these genes depends on coactivation by the cyclin E/CDK2-responsive p220NPAT protein (25, 28), and we found that the H4 genes that are most responsive to the HiNF-P·p220NPAT complex are also the most highly up-regulated at the G1/S phase transition. Furthermore, our study shows that there is a correlation between HiNF-P occupancy, responsiveness of the histone H4 promoter, and maximal induction at the G1/S phase transition. Taken together, these findings support the concept that the HiNF-P·p220NPAT complex is the principal regulatory module that functionally links the coordinate regulation of histone H4 genes with the onset of DNA replication at the G1/S phase transition.


   FOOTNOTES
 
* This work was supported by National Institutes of Health Grant GM32010 and the Deutsche Forschungsgemeinschaft. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

1 Both authors contributed equally to this work. Back

2 To whom correspondence should be addressed: Dept. of Cell Biology and Cancer Center, University of Massachusetts Medical School, 55 Lake Ave. North, Worcester, MA 01655. Tel.: 508-856-5625; Fax: 508-856-6800; E-mail: gary.stein{at}umassmed.edu.

3 The abbreviations used are: HiNF, histone nuclear factor; qPCR, quantitative PCR; ChIP, chromatin immunoprecipitation; qRT-PCR, quantitative reverse transcription-PCR; RPAs, RNase protection assays; PBS, phosphate-buffered saline; EMSA, electrophoretic mobility shift assay. Back


   ACKNOWLEDGMENTS
 
We thank all members of our laboratories as well as Wade Harper (Harvard Medical School) for thoughtful discussions and assistance with reagents. We also thank Judy Rask for expert editorial assistance.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

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